help button home button Endocrine Society Endocrinology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Endocrinology, doi:10.1210/en.2007-0864
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Iwashima, F.
Right arrow Articles by Hirata, Y.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Iwashima, F.
Right arrow Articles by Hirata, Y.
Endocrinology Vol. 149, No. 3 1009-1014
Copyright © 2008 by The Endocrine Society

Aldosterone Induces Superoxide Generation via Rac1 Activation in Endothelial Cells

Fumiko Iwashima, Takanobu Yoshimoto, Isao Minami, Maya Sakurada, Yuki Hirono and Yukio Hirata

Department of Clinical and Molecular Endocrinology, Tokyo Medical and Dental University Graduate School, Tokyo 113-8519, Japan

Address all correspondence and requests for reprints to: Takanobu Yoshimoto, M.D., Ph.D., Department of Clinical and Molecular Endocrinology, Tokyo Medical and Dental University Graduate School, 1-5-45 Yushima, Bunkyo-ku, Tokyo 113-8513, Japan. E-mail: tyoshimoto.cme{at}tmd.ac.jp.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Currently, aldosterone is believed to be involved in the development of cardiovascular injury as a potential cardiovascular risk hormone. However, its exact cellular mechanisms remain obscure. This study was undertaken to examine the effect of aldosterone on superoxide production in cultured rat aortic endothelial cells with possible involvement of the small GTP-binding (G) protein Rac1. The aldosterone levels showed a time-dependent (6–24 h) and dose-dependent (10–8 to 10–6 M) increase in superoxide generation, whose effect was abolished by mineralocorticoid receptor antagonist (eplerenone), Src inhibitor (PP2), and reduced nicotinamide adenine dinucleotide phosphate [NAD(P)H] oxidase inhibitor (apocynin). Aldosterone activated NADP(H) oxidase and Rac1, whose effects were abolished by eplerenone. The aldosterone-induced superoxide generation was abolished either by nonselective small G protein inhibitor (Clostridium difficile toxin A) or dominant-negative Rac1. Dominant-negative Rac1 also inhibited aldosterone-induced ACE gene expression. Thus, the present study is the first to demonstrate that aldosterone induces superoxide generation via mineralocorticoid receptor-mediated activation of NAD(P)H-oxidase and Rac1 in endothelial cells, thereby contributing to the development of aldosterone-induced vascular injury.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
ACCUMULATING LINES OF evidence suggest that the direct cardiovascular effects of aldosterone are responsible for inducing cardiovascular inflammation and fibrosis (aldosterone-induced vasculitis) (1, 2, 3). In addition, a series of clinical studies have demonstrated a distinct benefit of mineralocorticoid receptor (MR) antagonism in cardiovascular diseases (4, 5, 6).

There are several sources of reactive oxygen species (ROS) in mammalian cells, including mitochondrial electron transport system, xanthine oxidase, cyclooxygenase, nitric oxide synthase, and reduced nicotinamide adenine dinucleotide phosphate [NAD(P)H] oxidase (7). Among them, ROS derived from vascular NAD(P)H oxidase appears to be a major contributor for cardiovascular inflammation (7, 8, 9). Aldosterone has recently been shown to increase superoxide production and induces cardiovascular injury in mineralocorticoid-induced hypertensive animals, whose effects were blocked by the administration of subpressor doses of MR antagonists as well as antioxidants and/or NAD(P)H oxidase inhibitors (3, 10, 11, 12, 13). These experimental results suggest that oxidative stress induced by NAD(P)H oxidase plays an important role in the development of aldosterone-induced cardiovascular injury. However, the molecular mechanism underlying vascular superoxide generation and NAD(P)H oxidase activation by aldosterone remains unknown.

A small G protein, Rac, has been shown to be involved in the activation of NAD(P)H oxidase in phagocyte as well as in nonphagocyte cells, including cardiovascular cells (7, 14). Among several isoforms of Rac proteins, two isoforms (Rac1 and Rac2) play predominant roles in NAD(P)H oxidase activation (14). In contrast to the limited expression of Rac2 only in hematopoietic cells, Rac1 expressed ubiquitously is considered to be mainly responsible for NAD(P)H oxidase activation in nonhematopoietic cells (14).

It has been well recognized that endothelial damage is a primary event in the pathogenesis of cardiovascular diseases (15, 16). Previously, we reported that aldosterone directly acts on endothelial cells to induce angiotensin-converting enzyme and osteopontin expression in vitro (17, 18). In addition, we have recently reported marked increase in oxidant stress in the endothelium from aldosterone-induced hypertensive rats, the effect of which was eliminated by the selective MR antagonist eplerenone or the superoxide dismutase mimetic tempol (11). These findings postulate that aldosterone possibly acts directly on endothelial cells to induce oxidative stress and vascular inflammation. However, the mechanism of the aldosterone effect on superoxide generation in endothelial cells remains unknown.

The present study aims to examine whether aldosterone directly stimulates NAD(P)H oxidase to generate superoxide in cultured rat aortic endothelial cells (RAECs) and, if so, to determine its molecular mechanism, particularly the potential involvement of the small G protein Rac1 in NAD(P)H oxidase activation.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Aldosterone was purchased from Acros Organics (Geel, Belgium); dihydroethidium (DHE) from Invitrogen (Carlsbad, CA); rabbit anti-actin polyclonal antibody, EGTA, phenylmethylsulfonyl fluoride (PMSF), N,N'-dimethyl-9,9'-biacridinium dinitrate (lucigenin), and diphenyleneiodonium chloride (DPI) from Sigma (St. Louis, MO); apocynin, Clostridium difficile (CD) toxin A, and PP2 from EMD Biosciences, Inc. (San Diego, CA); Matrigel and endothelial cell growth supplement from BD Biosciences (Bedford, MA); mouse anti-Rac1 monoclonal antibody from Upstate Biotechnology Inc. (Lake Placid, NY); anti-G6PD antibody from Bethyl Laboratories Inc. (Montgomery, TX); Rac1N17 cDNA from the University of Missouri-Rolla cDNA Resource Center (Rolla, MO); and LacZ cDNA from Promega (Madison, WI). Eplerenone was kindly provided by Pfizer Inc. (New York, NY). The adenovirus shuttle plasmid pAdCMV295MCSloxP (19) and the sub360 adenovirus cosmid cS360loxP (19) were generous gifts from Elizabeth G. Nabel (National Heart, Lung, and Blood Institute, Bethesda, MD). Aldosterone and eplerenone were dissolved in dimethylsulfoxide (DMSO) at 10–2 M in stock solution.

Cell culture
RAECs were prepared from the thoracic aorta of 6-wk-old male Sprague Dawley rats fed with standard chow using the explants method as described previously (17, 18, 20). Briefly, thoracic aorta was sterilely removed from a rat deeply anesthetized by an ip injection of sodium pentobarbital. The vessel was gently cleaned in Hanks’ balanced salt solution and cut into flat segments of approximately 4 mm2. The flat aortic segments were placed endothelial-side down on Matrigel equilibrated with growth medium (medium 199 containing 10% fetal bovine serum and 15 µg/ml endothelial cell growth supplement) in a 35-mm dish. The aortic explants were removed after 4–8 d, depending on the degree of outgrowth of endothelial cells. Cells on Matrigel were passaged with 0.2% collagenase in Hanks’ balanced salt solution and replated in growth medium onto a 60-mm type 1 collagen-coated dish. These RAECs were routinely subcultured by treatment with 0.05% trypsin and 0.02% EDTA. Characterization of isolated cells as endothelial cells was described previously (18). Subcultured RAECs (fourth to seventh passages) were starved with medium 199 containing 0.5% calf serum (starvation medium) for 24 h and used for subsequent experiments. All of the experiments were conducted in starvation medium containing 0.1% DMSO; the medium containing 0.1% DMSO alone served as vehicle control.

Because the responses to aldosterone greatly varied depending on whether the cells were from different passage numbers and/or from the different rats (batch difference), cells from the same passage number derived from one animal were used for a series of each experiment to minimize the batch differences.

Measurement of intracellular superoxide anion levels
The intracellular superoxide anion level was measured using the cell-permeable dye DHE as previously described (21). In brief, 1 x 105 cells seeded and cultured on a 35-mm glass-bottom dish (Matsunami, Tokyo, Japan) for 24 h were subjected to serum starvation; 80–90% confluent cells after 24 h starvation were treated with or without the test compound for the indicated duration. Cells were then incubated with 5 µM DHE for 20 min at 37 C, washed in Krebs-Ringer phosphate buffer (15.9 mM NaH2PO4, 11 mM glucose, 1.2 mM MgSO4, 0.54 mM CaCl2, 4.8 mM KCl, and 120 mM NaCl) (pH 7.3), and imaged by inverted fluorescent microscopy under conditions of 514 nm excitation and 580 ± 30 nm band pass filter (IX71; Olympus, Tokyo, Japan). Data analysis was performed as previously described (22, 23).

Measurement of NAP(P)H oxidase activity
NAD(P)H oxidase activity was measured by the method described previously (23) with minor modification. Briefly, starved cells grown on 10-cm dishes were pretreated with or without eplerenone (3 x 10–7 M) for 1 h and stimulated with or without aldosterone (10–8 M) for 12 h. After completion, cells were harvested, resuspended in lysis buffer composed of 5 x 10–2 M phosphate buffer (pH 7.0), containing 10–3 M EGTA, 10 µg/ml aprotinin, and 10–3 M PMSF, and dounced on ice. An aliquot (50 µg protein) of the lysate was brought into assay buffer composed of 5 x 10–2 M phosphate buffer (pH 7.0) containing 10–3 M EGTA, 1.5 x 10–1 M sucrose, and 1 x 10–5 M dark-adapted lucigenin with or without 1 x 10–4 M DPI in a 96-well white assay plate (Corning, Rochester, NY). After preincubation at 37 C for 5 min, NADH (10–4 M) was added, and luminescence was measured with a microplate luminometer (LB96V; Berthold Japan, Tokyo, Japan) over 1-min intervals for a total 15 min. NAD(P)H oxidase activity was measured as NAD(P)H oxidase-dependent superoxide anion (O2–) production by integrated lucigenin chemiluminescence over 15 min without DPI subtracted from that with DPI.

Measurement of activated Rac1
The GTP-bound form of active Rac1 was measured using the Rac activation assay kit (Upstate Biotechnology) according to the affinity purification method by using the p21-activated kinase-1 (PAK-1) protein-binding domain peptide (24). In brief, cells grown on 6-cm dishes were lysed with 250 µl ice-cold lysis buffer [25 mM HEPES, 150 nM NaCl, 1% IGEPAL CA-630, 10 mM MgCl2, 10% glycerol, 1 mM EDTA, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 1 mM NaF, and 1 mM NaVO4 (pH 7.5)]. Each 25 µl (10%) lysate was maintained for the subsequent immunoblot analysis for total Rac1 and actin, and the remaining lysates were incubated using 8 µg PAK-1 protein-binding domain agarose for 1 h. After extensive washing with the lysis buffer, the agarose was boiled in Laemmli sample buffer. The affinity-purified samples or aliquots containing total cell lysates were resolved using 15% SDS-PAGE and analyzed by immunoblot analysis using the anti-Rac1 antibody or anti-actin antibody, as previously described (22).

Gene transfer of the adenovirus vector
Replication-defective adenovirus encoding the dominant-negative form of Rac1 (RacN17) (25) or β-galactosidase (LacZ) was prepared by Cre/loxP-mediated recombination between the sub3360 adenoviral cosmid (cS360loxP) and adenovirus shuttle plasmid (pAdCMV2295MCSloxP) inserted with each cDNA expression cassette (19). Changing the amino acid residue from Ser to Asn at the position of 17 of Rac1 results in preferential affinity for GDP rather than GTP and behaves as dominant-negative mutant (26). After confirmation of protein expression, each recombinant adenovirus was propagated in the HEK293 cells and purified by using cesium chloride gradient ultracentrifugation. Viral titers were determined by using a plaque assay methods.

With regard to the gene transfer study, RAECs were infected with 50 plaque-forming units virus per cell (multiplicity of infection, 50) in medium 199 containing 2% fetal calf serum for 2 h, followed by a 48-h incubation in medium 199 containing 10% fetal calf serum and starvation before the experiment.

Quantification of mRNA
Steady-state mRNA levels of rat ACE and osteopontin were quantified with a real-time quantitative RT-PCR using fluorescent SYBR Green technology (LightCycler; Roche Molecular Biochemicals, Mannheim, Germany) as described previously (17, 18). Rat acid ribosomal phosphoprotein P0 (ARPP P0) mRNA levels were quantified by TaqMan fluorescence methods as described previously (17, 18), except using QuantiTect Probe PCR Kit (QIAGEN, Valencia, CA) and LightCycler. Total RNA was extracted, first-strand cDNA synthesized, and amplification reaction performed as described (17, 18). PCR primers, TaqMan probes, and size of each PCR product are as follows: ACE, forward 5'-CAGAGGCCAACTGGCATTAT-3' and reverse 5'-CTGGAAGTTGCT CACGTCAA-3', product size 137 bp; osteopontin, forward 5'-AGTGGTTTGCCTTTGCCTGTT-3' and reverse 5'-TCAGCCAAGTGGCTACAGCAT-3', product size 122 bp; ARPP P0 forward 5'-TAGAGGGTGTCCGCAATGTG-3', reverse 5'-GACAAAGCCAGGACCCTTTTGT-3', TaqMan probe 5'-ACCCGACTGTTGCCTCAGTGCCTCACTCCA-3', product size 107 bp.

Statistical analysis
Data are expressed as mean ± SEM. The differences between groups were examined for statistical significance using the t test or ANOVA with Dunn’s post hoc test, if appropriate. P values < 0.05 were considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
We first examined the effect of aldosterone on intracellular superoxide generation in RAECs by measuring DHE fluorescence. Aldosterone (10–8 M) increased intracellular superoxide generation in a time-dependent manner (6–24 h); a significant increase was observed as early as 6 h, and it peaked at 12 h (Fig. 1AGo). Incubation with vehicle (0.1% DMSO) alone by 24 h did not affect intracellular ROS levels (Fig. 1AGo) or cell viability (data not shown). Treatment with aldosterone for 12 h increased intracellular superoxide levels in a dose-dependent manner (10–8 to 10–6 M); a significant increase was induced at a concentration as low as 10–8 M and the peak at 10–6 M (Fig. 1BGo).


Figure 1
View larger version (16K):
[in this window]
[in a new window]

  		FIG. 1. Aldosterone induces superoxide generation in RAECs. RAECs were incubated with aldosterone (10–8 M) for the indicated times (A) or incubated with aldosterone at the indicated concentration for 12 h (B). Intracellular superoxide generation was measured using DHE fluorescence. The fluorescence images of the cells were recorded randomly at three different fields; the mean fluorescence intensities of 150 cells in one dish were calculated as one experiment. Each circle represents the mean ± SEM of three independent experiments; the values are expressed as fold increase over the control. Open and closed circles indicate treatment with vehicle (0.1% DMSO) or aldosterone, respectively. *, P < 0.05 vs. control.

 
The aldosterone-induced superoxide generation was blocked by pretreatment with the selective MR antagonist eplerenone (3 x 10–7 M) (Fig. 2AGo) and the NAD(P)H oxidase inhibitor apocynin (3 x 10–4 M) (Fig. 2BGo). Therefore, aldosterone-induced superoxide generation results from MR-mediated NAD(P)H oxidase activation in endothelial cells.


Figure 2
View larger version (22K):
[in this window]
[in a new window]

  		FIG. 2. Aldosterone-induced superoxide generation was blocked by MR antagonist, NAD(P)H oxidase inhibitor, small G protein inhibitor, and Src inhibitor in RAECs. After pretreatment for 1 h with or without eplerenone (3 x 10–7 M) (A), apocynin (3 x 10–4 M) (B), CD toxin A (10 ng/ml) (C), or PP2 (10–5 M) (D), cells were incubated with aldosterone (10–8 M) or vehicle for 12 h. Intracellular superoxide generation was measured using DHE fluorescence. Each column shows fold increase over the control and is the mean of three independent experiments. Closed and open columns indicate with and without aldosterone treatment, respectively. *, P < 0.05 vs. control; {dagger}, P < 0.05 vs. aldosterone without inhibitor.

 
Because Rac, a small G protein, has been shown to play a crucial role in NAD(P)H oxidase activation and superoxide generation (14), we examined the possible involvement of Rac in aldosterone-induced superoxide generation in RAECs using CD toxin A, a nonselective inhibitor of the small G proteins (Rho, Rac, and cdc42). CD toxin A (10 ng/ml) completely inhibited the aldosterone-induced superoxide generation (Fig. 2CGo).

Because it has been shown that aldosterone activates NAD(P)H oxidase via Src in cultured rat vascular smooth muscle cells (VSMCs) (27), we tested the effect of PP2, a selective Src inhibitor, on aldosterone-induced superoxide generation. PP2 (10–5 M) completely inhibited the aldosterone-induced superoxide generation (Fig. 2DGo).

To confirm that the aldosterone-indcued superoxide generation was derived from NAD(P)H oxidase, we measured NAD(P)H oxidase activity using a lucigenin chemiluminescence assay. Treatment with aldosterone (10–8 M) showed a significant increase in NAD(P)H oxidase activity, whose effect was inhibited by eplerenone (Fig. 3Go).


Figure 3
View larger version (16K):
[in this window]
[in a new window]

  		FIG. 3. Aldosterone induces NAD(P)H oxidase activation via MR in RAECs. After pretreatment with or without eplerenone (3 x 10–7 M) for 1 h, RAECs were incubated with aldosterone (10–8 M) for 12 h. NAD(P)H oxidase activity was determined by the lucigenin chemiluminescence assay as described in Materials and Methods. Each column shows fold increase over control and is the mean of five independent experiments. Closed and open columns indicate with and without aldosterone treatment, respectively. *, P < 0.05 vs. control; {dagger}, P < 0.05 vs. aldosterone without eplerenone.

 
We then investigated whether aldosterone activates Rac1 in endothelial cells by measuring the levels of GTP-bound active Rac1. Aldosterone significantly (P < 0.05) increased the GTP-bound Rac1 levels in a time-dependent manner (6–24 h) identical to that observed for superoxide generation, without affecting the total Rac1 protein levels (Fig. 4Go, A and B). Pretreatment with eplerenone abolished the aldosterone-induced Rac1 activation (Fig. 4Go, C and D), suggesting that aldosterone activates Rac1 via MR.


Figure 4
View larger version (37K):
[in this window]
[in a new window]

  		FIG. 4. Aldosterone induces Rac1 activation via MR in RAECs. After pretreatment with or without eplerenone (10–5 M) for 1 h, RAECs were incubated with aldosterone (10–8 M) for the indicated times (A and B) and for 12 h (C and D). Rac activity was measured using PAK-1 affinity purification; total Rac1 and actin were detected by immunoblot analysis as described in Materials and Methods. A and C, Representative blots with GTP-Rac1 (top), total-Rac1 (middle), and actin (bottom); B and D, the ratio of GTP-Rac1 to total Rac1 signals was quantified by densitometric scanning; each column shows fold increase over control and is the mean of three independent experiments. Closed and open columns indicate with and without aldosterone treatment, respectively. *, P < 0.05 vs. control; {dagger}, P < 0.05 vs. aldosterone without eplerenone.

 
To further confirm the specific role of Rac1 in aldosterone-induced superoxide generation in RAECs, the cells were subjected to gene transfer by using the recombinant dominant-negative Rac1 adenovirus (Ad-RacN17) and LacZ adenovirus (Ad-LacZ) as the control. As shown in Fig. 5Go, gene transfer using Ad-RacN17 completely blocked the aldosterone-induced superoxide generation, whereas gene transfer with Ad-LacZ had no effect.


Figure 5
View larger version (13K):
[in this window]
[in a new window]

  		FIG. 5. Aldosterone-induced superoxide generation is blocked by adenovirus gene transfer of dominant-negative Rac1 (RacN17) in RAECs. RAECs were infected with Ad-LacZ or Ad-RacN17. RAECs treated without the virus were used as a mock-infection control. After 48 h, the cells were incubated with or without aldosterone (10–8 M) for 12 h. DHE fluorescence was measured, and the data were calculated and plotted as described in Fig. 2Go. *, P < 0.05. NS, Not significant.

 
Because aldosterone increases ACE and osteopontin gene expression in RAECs (17, 18), we examined whether dominant-negative Rac1 affects ACE and osteopontin mRNA expression in response to aldosterone (10–8 M). Gene transfer with Ad-RacN17 completely blocked the aldosterone-induced ACE mRNA expression, whereas that with Ad-LacZ was without effect (Fig. 6Go). By contrast, gene transfer with either Ad-RacN17 or Ad-LacZ alone caused a marked decrease in osteopontin mRNA expression even under basal conditions, possibly due to a nonspecific effect by adenoviral particles (data not shown).


Figure 6
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 6. Aldosterone-induced ACE gene expression is blocked by adenovirus gene transfer of RacN17 in RAECs. RAECs were infected with Ad-LacZ or Ad-RacN17; cells treated without the virus were used as a mock-infection control. After 48 h, the cells were incubated with or without aldosterone (10–8 M) for 18 h; ACE mRNA levels were measured by real-time RT-PCR. Each column shows fold increase over control and is the mean from five independent experiments. *, P < 0.05. NS, Not significant.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The present study demonstrated for the first time that aldosterone induces superoxide generation via NAD(P)H oxidase and Rac1 activation through MR in RAECs. Aldosterone-induced superoxide generation was completely blocked by a nonselective small G protein inhibitor and adenoviral gene transfer of the dominant-negative Rac1, suggesting that aldosterone-induced Rac1 activation plays a crucial role in superoxide generation due to NAD(P)H oxidase in endothelial cells.

Recently, numerous experimental studies have revealed the potential direct role of aldosterone as a cardiovascular risk hormone in the development of cardiovascular disease via oxidative stress (1, 2, 3). There are several sources of ROS in mammalian cells, including mitochondrial electron transport system, xanthine oxidase, cyclooxygenase, nitric oxide synthase, and NAD(P)H oxidase. Because agonist-stimulated superoxide generation in cardiovascular cells is inhibited by dominant-negative Rac1 and antisense oligonucleotides for NAD(P)H oxidase components, NAD(P)H oxidase-derived ROS appears to be largely responsible for the development of cardiovascular injury (7). In accordance with this, several recent studies have revealed that increased superoxide generation by NAD(P)H oxidase is involved in aldosterone-induced oxidative stress and inflammatory response in cardiovascular tissue, including cardiomyocytes and VSMCs (3, 10, 11, 12, 13). We have recently reported that redox-sensitive proinflammatory gene expression and endothelial oxidative stress are hallmarks of vascular injury in the aldosterone-induced hypertensive rat model (11). In the present study, we clearly demonstrated that the aldosterone-MR pathway stimulates superoxide generation via NAD(P)H oxidase in vascular endothelial cells. This result is almost in agreement with a recent report stating that aldosterone stimulates the formation of ROS in human umbilical vein endothelial cells (28). Taken together with the previous data that aldosterone induces NAD(P)H oxidase activation in VSMCs (27), the present data of aldosterone-induced NAD(P)H oxidase activation in endothelial cells lend support to the contention that oxidative stress derived from vascular NAD(P)H oxidase activation is largely responsible for the development of aldosterone-induced cardiovascular injury.

The cellular mechanism of NAD(P)H oxidase activation in phagocytes such as neutrophils and macrophages has been well characterized (8, 14, 29); activation of the small G protein Rac and phosphorylation of the p47phox trigger assembly comprising membrane (Nox and p22phox) and cytosolic components (p47phox, p67phox, p40phox, and Rac2) of NAD(P)H oxidase, leading to its complete activation. However, the mode of NAD(P)H oxidase activation by aldosterone in vascular cells has not been well characterized. Because Rac1 is a predominant Rac subtype in nonphagocyte cells (14), we examined the effect of aldosterone on the levels of GTP-bound (active) Rac1 in endothelial cells. In the present study, we demonstrated for the first time that aldosterone increased the GTP-bound form of Rac1 via MR in the same time as that required for superoxide generation in RAECs. Furthermore, we also demonstrated that Rac1 activation is indispensable for aldosterone-induced superoxide generation; the abovementioned result is based on the observations that inhibition of Rac by treatment with either CD toxin A or gene transfer of dominant-negative Rac1 abolished aldosterone-induced superoxide generation. In the present study, we also found that a src inhibitor, PP2, blocked the aldosterone-induced superoxide generation in RAECs, suggesting the possible involvement of Src in NAD(P)H oxidase activation. These findings complement the previous findings that aldosterone activates NAD(P)H oxidase via Src in VSMCs (27). Because Rac1 activation is essential for the angiotensin II-mediated NA(D)PH oxidase activation with possible involvement of Src as an upstream signaling molecule in VSMCs (30), it appears that angiotensin II and aldosterone share the Src-Rac pathway in common for NAD(P)H oxidase activation in vascular cells.

It is an important issue as to whether aldosterone-induced superoxide generation is via nongenomic or classical genomic action. Unfortunately, we could not obtain the conclusive evidence, because various doses of actinomycin D (1–10 µM) and cycloheximide (5–20 µg/ml) inhibited basal ROS generation by approximately 50% in RAECs, suggesting requirement of de novo gene expression for ROS generation in RAECs under the steady-state conditions (unpublished data). Nevertheless, genomic mechanism(s) for aldosterone on Rac1 activation seems likely because Rac1 activation and superoxide generation by aldosterone took longer times (at least 6 h).

We have reported that ACE gene expression by aldosterone was increased in RAECs in vitro (18) as well as in cardiovascular tissue in vivo (11). In the present study, gene transfer of dominant-negative Rac1 abolished the aldosterone-induced ACE gene expression in RAECs, suggesting the possible involvement of Rac1 activation by aldosterone in ACE gene expression via a superoxide-dependent mechanism. In contrast, we have recently shown that treatment with tempol, a superoxide dismutase mimetic, did not affect the up-regulated ACE gene expression in aortic tissue from aldosterone-induced hypertensive rats (11). Therefore, it is suggested that a diversity of molecular signaling mechanisms other than a redox-sensitive pathway are involved in ACE gene expression in cardiovascular tissue of aldosterone-induced hypertension.

Recently, it has also been demonstrated that aldosterone decreased the expression and activity of glucose-6-phosphate dehydrogenase (G6PD) to reduce glutathione (GSH)-mediated redox regulation in bovine aortic endothelial cells after long-term (24 h) incubation, resulting in increased ROS levels and decreased NO levels (31). Consumption of GSH via increased NAD(P)H oxidase-derived superoxide and impaired GSH production due to decreased G6PD activity might contribute to the intracellular oxidative state and uncoupled endothelial NO synthase, which generates superoxide rather than NO (32). Because aldosterone decreased G6PD expression and G6PD activities in endothelial cells and aortic tissue from C3H mice (31), we examined whether aldosterone affects G6PD expression in RAECs. However, treatment with aldosterone (10–9 to 10–7 M) for 24 h did not cause any changes in G6PD protein levels in RAECs (supplemental figure, published as supplemental data on The Endocrine Society’s Journals Online web site at http://endo.endojournals.org). Such a discrepancy might be accounted for by the different experimental conditions or species difference of endothelial cells examined. In contrast, the present study revealed that MR-mediated superoxide generation was activated by aldosterone via Rac1-dependent NAD(P)H oxidase in RAECs. Nevertheless, it is reasonable to speculate that aldosterone may use diverse signaling pathways in endothelial cells, including NAD(P)H oxidase activation and/or reduction in GSH-mediated redox regulation, thus constituting a vicious cycle for acceleration of endothelial dysfunction via oxidative stress.

In conclusion, our present study demonstrates that aldosterone induces superoxide generation via MR-mediated activation of NAD(P)H oxidase and Rac1 in vascular endothelial cells, thereby contributing to the development of aldosterone-induced vascular injury.


   Acknowledgments
 
We thank Dr. Elizabeth G. Nabel for generously gifting us with pAdCMV295MCSloxP and cS360loxP.


   Footnotes
 
This study was supported in part by a grant-in-aid from the Ministry of Education, Science, Sports, and Culture and the Ministry of Health, Welfare, and Labor of Japan.

Disclosure Statement: The authors have nothing to disclose.

First Published Online December 13, 2007

Abbreviations: CD, Clostridium difficile; DHE, dihydroethidium; DMSO, dimethylsulfoxide; DPI, diphenyleneiodonium chloride; G6PD, glucose-6-phosphate dehydrogenase; GSH, glutathione; MR, mineralocorticoid receptor; NAD(P)H, reduced nicotinamide adenine dinucleotide phosphate; PAK-1, p21-activated kinase-1; PMSF, phenylmethylsulfonyl fluoride; RAEC, rat aortic endothelial cell; ROS, reactive oxygen species; VSMC, vascular smooth muscle cell.

Received June 27, 2007.

Accepted for publication November 30, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Funder JW 2004 Aldosterone, mineralocorticoid receptors and vascular inflammation. Mol Cell Endocrinol 217:263–269[CrossRef][Medline]
  2. Rocha R, Funder JW 2002 The pathophysiology of aldosterone in the cardiovascular system. Ann NY Acad Sci 970:89–100[Medline]
  3. Schiffrin EL 2006 Effects of aldosterone on the vasculature. Hypertension 47:312–318[Free Full Text]
  4. Pitt B, Reichek N, Willenbrock R, Zannad F, Phillips RA, Roniker B, Kleiman J, Krause S, Burns D, Williams GH 2003 Effects of eplerenone, enalapril, and eplerenone/enalapril in patients with essential hypertension and left ventricular hypertrophy: the 4E-left ventricular hypertrophy study. Circulation 108:1831–1838[Abstract/Free Full Text]
  5. Pitt B, Remme W, Zannad F, Neaton J, Martinez F, Roniker B, Bittman R, Hurley S, Kleiman J, Gatlin M, Pitt B, Zannad F, Remme WJ, Cody R, Castaigne A, Perez A, Palensky J, Wittes J 2003 Eplerenone, a selective aldosterone blocker, in patients with left ventricular dysfunction after myocardial infarction. N Engl J Med 348:1309–1321[Abstract/Free Full Text]
  6. Pitt B, Zannad F, Remme WJ, Cody R, Castaigne A, Perez A, Palensky J, Wittes J 1999 The effect of spironolactone on morbidity and mortality in patients with severe heart failure. Randomized Aldactone Evaluation Study Investigators. N Engl J Med 341:709–717[Abstract/Free Full Text]
  7. Ushio-Fukai M, Alexander RW 2004 Reactive oxygen species as mediators of angiogenesis signaling: role of NAD(P)H oxidase. Mol Cell Biochem 264:85–97[CrossRef][Medline]
  8. Lassegue B, Clempus RE 2003 Vascular NAD(P)H oxidases: specific features, expression, and regulation. Am J Physiol Regul Integr Comp Physiol 285:R277–R297
  9. Griendling KK, Sorescu D, Ushio-Fukai M 2000 NAD(P)H oxidase: role in cardiovascular biology and disease. Circ Res 86:494–501[Abstract/Free Full Text]
  10. Beswick RA, Dorrance AM, Leite R, Webb RC 2001 NADH/NADPH oxidase and enhanced superoxide production in the mineralocorticoid hypertensive rat. Hypertension 38:1107–1111[Abstract/Free Full Text]
  11. Hirono Y, Yoshimoto T, Suzuki N, Sugiyama T, Sakurada M, Takai S, Kobayashi N, Shichiri M, Hirata Y 2007 Angiotensin II receptor type 1-mediated vascular oxidative stress and proinflammatory gene expression in aldosterone-induced hypertension: the possible role of local renin-angiotensin system. Endocrinology 148:1688–1696[Abstract/Free Full Text]
  12. Rudolph AE, Rocha R, McMahon EG 2004 Aldosterone target organ protection by eplerenone. Mol Cell Endocrinol 217:229–238[CrossRef][Medline]
  13. Sun Y, Zhang J, Lu L, Chen SS, Quinn MT, Weber KT 2002 Aldosterone-induced inflammation in the rat heart: role of oxidative stress. Am J Pathol 161:1773–1781[Abstract/Free Full Text]
  14. Hordijk PL 2006 Regulation of NADPH oxidases: the role of Rac proteins. Circ Res 98:453–462[Abstract/Free Full Text]
  15. Davignon J, Ganz P 2004 Role of endothelial dysfunction in atherosclerosis. Circulation 109(23 Suppl 1):III27–III32
  16. Ross R 1999 Atherosclerosis: an inflammatory disease. N Engl J Med 340:115–126[Free Full Text]
  17. Sugiyama T, Yoshimoto T, Hirono Y, Suzuki N, Sakurada M, Tsuchiya K, Minami I, Iwashima F, Sakai H, Tateno T, Sato R, Hirata Y 2005 Aldosterone increases osteopontin gene expression in rat endothelial cells. Biochem Biophys Res Commun 336:163–167[CrossRef][Medline]
  18. Sugiyama T, Yoshimoto T, Tsuchiya K, Gochou N, Hirono Y, Tateno T, Fukai N, Shichiri M, Hirata Y 2005 Aldosterone induces angiotensin converting enzyme gene expression via a JAK2-dependent pathway in rat endothelial cells. Endocrinology 146:3900–3906[Abstract/Free Full Text]
  19. Aoki K, Barker C, Danthinne X, Imperiale MJ, Nabel GJ 1999 Efficient generation of recombinant adenoviral vectors by Cre-lox recombination in vitro. Mol Med 5:224–231[Medline]
  20. McGuire PG, Orkin RW 1987 Isolation of rat aortic endothelial cells by primary explant techniques and their phenotypic modulation by defined substrata. Lab Invest 57:94–105[Medline]
  21. Brandes RP, Miller FJ, Beer S, Haendeler J, Hoffmann J, Ha T, Holland SM, Gorlach A, Busse R 2002 The vascular NADPH oxidase subunit p47phox is involved in redox-mediated gene expression. Free Radic Biol Med 32:1116–1122[CrossRef][Medline]
  22. Yoshimoto T, Fukai N, Sato R, Sugiyama T, Ozawa N, Shichiri M, Hirata Y 2004 Antioxidant effect of adrenomedullin on angiotensin II-induced reactive oxygen species generation in vascular smooth muscle cells. Endocrinology 145:3331–3337[Abstract/Free Full Text]
  23. Yoshimoto T, Gochou N, Fukai N, Sugiyama T, Shichiri M, Hirata Y 2005 Adrenomedullin inhibits angiotensin II-induced oxidative stress and gene expression in rat endothelial cells. Hypertens Res 28:165–172[CrossRef][Medline]
  24. Benard V, Bohl BP, Bokoch GM 1999 Characterization of rac and cdc42 activation in chemoattractant-stimulated human neutrophils using a novel assay for active GTPases. J Biol Chem 274:13198–13204[Abstract/Free Full Text]
  25. Nobes CD, Hall A 1995 Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81:53–62[CrossRef][Medline]
  26. Ridley AJ, Paterson HF, Johnston CL, Diekmann D, Hall A 1992 The small GTP-binding protein rac regulates growth factor-induced membrane ruffling. Cell 70:401–410[CrossRef][Medline]
  27. Callera GE, Touyz RM, Tostes RC, Yogi A, He Y, Malkinson S, Schiffrin EL 2005 Aldosterone activates vascular p38MAP kinase and NADPH oxidase via c-Src. Hypertension 45:773–779[Abstract/Free Full Text]
  28. Nagata D, Takahashi M, Sawai K, Tagami T, Usui T, Shimatsu A, Hirata Y, Naruse M 2006 Molecular mechanism of the inhibitory effect of aldosterone on endothelial NO synthase activity. Hypertension 48:165–171[Abstract/Free Full Text]
  29. Babior BM, Lambeth JD, Nauseef W 2002 The neutrophil NADPH oxidase. Arch Biochem Biophys 397:342–344[CrossRef][Medline]
  30. Seshiah PN, Weber DS, Rocic P, Valppu L, Taniyama Y, Griendling KK 2002 Angiotensin II stimulation of NAD(P)H oxidase activity: upstream mediators. Circ Res 91:406–413[Abstract/Free Full Text]
  31. Leopold JA, Dam A, Maron BA, Scribner AW, Liao R, Handy DE, Stanton RC, Pitt B, Loscalzo J 2007 Aldosterone impairs vascular reactivity by decreasing glucose-6-phosphate dehydrogenase activity. Nat Med 13:189–197[CrossRef][Medline]
  32. Vasquez-Vivar J, Kalyanaraman B, Martasek P 2003 The role of tetrahydrobiopterin in superoxide generation from eNOS: enzymology and physiological implications. Free Radic Res 37:121–127[CrossRef][Medline]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Iwashima, F.
Right arrow Articles by Hirata, Y.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Iwashima, F.
Right arrow Articles by Hirata, Y.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Endocrinology Endocrine Reviews J. Clin. End. & Metab.
Molecular Endocrinology Recent Prog. Horm. Res. All Endocrine Journals